X.B. is supported by the National Natural Science Foundation of China (32371287), the Fundamental Research Funds for the Central Universities (63223043 and 63233053), and the Talent Training Project at Nankai University (035-BB042112). Y.L. is supported by the National Natural Science Foundation of China (12022409 and T2221001) and the CAS Key Research Program of Frontier Sciences (ZDBS-LY-SLH015). L.M. is supported by the National Natural Science Foundation of China (32271274 and 31770812).

The details of the reagents and equipment used in this study are listed in the Table of Materials.
1. Preparation of modified coverslip for protein immobilization chamber
<ol>
	<li>Preparation
	<ol>
		<li>Prepare 50 mL of piranha solution (compose of concentrated sulfuric acid and hydrogen peroxide in a ratio of 3:1) by slowly adding hydrogen peroxide to concentrated sulfuric acid to prevent excessive temperatures. Place it carefully in a fume hood.</li>
		<li>Prepare sodium ethoxide solution. Dissolve 2 g of NaOH in 15 mL of ddH2O, add 35 mL of absolute ethanol, and mix completely. Leave at room temperature.</li>
		<li>Prepare high-salt solution (0.1 M of NaHCO3 and 0.6 M of K2SO4). Dissolve completely and aliquot 500 &#181;L/tube. Store at -20 &#176;C.</li>
		<li>Aliquot SVA-mPEG and SVA-mPEG-biotin into approximately 30 mg per tube and 3 mg per tube. Store at -20 &#176;C.</li>
		<li>Customize microscope slides (75 mm &#215; 26 mm &#215; 1 mm) with 6 pairs of 1.2-mm-diameter holes.</li>
	</ol>
	</li>
	<li>Cleaning the surface of the coverslip
	<ol>
		<li>Use tweezers to pick up eight coverslips and place them in a staining jar. Add acetone (~50 mL) to cover the coverslips, place the staining jar in an ultrasonic cleaner for 30 min, and then rinse the coverslips three times with ddH2O.</li>
		<li>Wash the coverslips with methanol as in step 1.2.1.</li>
		<li>Add the piranha solution (step 1.1.1) to the staining jar and heat it in a 95 &#176;C water bath kettle for 2 h. After the staining jar has cooled to room temperature, rinse the coverslips at least six times with ddH2O.</li>
		<li>Add the sodium ethoxide solution into the staining jar, place it in an ultrasonic cleaner for 15 min, and then rinse three times with ddH2O.</li>
		<li>Add ddH2O to the staining jar to cover the coverslips, place in an ultrasonic cleaner for 15 min, and subsequently rinse 3 times with ddH2O.</li>
	</ol>
	</li>
	<li>Aminosilanization modification of the coverslip surface.
	<ol>
		<li>Clamp the coverslips in the staining jar on a clean bench, dry them with nitrogen, and put them into another staining jar (dried in advance).</li>
		<li>Bake the staining jar in a drying oven at 120 &#176;C for 30 min, and then cool to room temperature in a desiccator.</li>
		<li>Add 47.5 mL of methanol, 2.5 mL of acetic acid, and 0.5 mL of 3-Aminopropyltriethoxysilane (APTES) to the beaker (dried in advance), mix evenly, add to the staining jar, and incubate for 10 min.</li>
		<li>Rinse the coverslips in the staining jar with ddH2O at least three times and sonicate for 5 min.</li>
		<li>Take out the coverslips one by one, rinse with ddH2O, blow dry with nitrogen, and put them into a 10-cm-dimeter Petri dish to prepare for PEG modification. 
		NOTE: APTES modification must be carried out in a dry environment.</li>
	</ol>
	</li>
	<li>Modification of coverslip surface with SVA-mPEG-Biotin and SVA-mPEG 
	NOTE: Biotin modification of the coverslip surface helps retain biotinylated proteins (or proteins binding to biotin-anti-His) via streptavidin.
	<ol>
		<li>Dissolve the packaged SVA-mPEG and SVA-mPEG-Biotin in the high-salt solution (1 mg corresponding to 10 &#181;L high-salt solution) completely. Add the SVA-mPEG-Biotin solution to the SVA-mPEG solution at a ratio of 1:100. 
		NOTE: Use a vortex to shake the solution to ensure that SVA-mPEG or SVA-mPEG-Biotin is completely dissolved. The SVA-mPEG or SVA-mPEG-Biotin aqueous solution should be prepared before use.</li>
		<li>Drop the prepared mix solution on a coverslip and cover it with another coverslip. This process should prevent the generation of bubbles. Incubate coverslips fully with appropriate humidity for more than 2 h or overnight.</li>
		<li>Separate the coverslips after modification is completed, rinse them with ddH2O, and blow dry with nitrogen. Mark the surface of the coverslip that has not been modified with SVA-mPEG and SVA-mPEG-biotin.</li>
		<li>Carefully place the modified coverslip in a 50 mL microcentrifuge tube. Store at -20 &#176;C after vacuuming. 
		NOTE: The modified coverslip can be stored under vacuum for 1 month.</li>
	</ol>
	</li>
</ol>
2. Preparation of microscope slides for protein immobilization chamber
<ol>
	<li>Wash the microscope slides as described for coverslips in step 1.2.</li>
	<li>Blow the microscope slides dry with nitrogen and put them into 50 mL microcentrifuge tubes.</li>
	<li>Store the microscope slides as described in step 1.4.4.</li>
</ol>
3. Protein expression and purification
<ol>
	<li>Preparation
	<ol>
		<li>Plasmids: Clone atlastin family member of interest or the pathogenic mutation of interest into a bacterial expression vector, for example, pET-28a, and add an Avitag27. Introduce cysteines at relevant positions by site-directed mutagenesis (for example, T51C and K400C27).</li>
		<li>Prepare the following buffers.
		<ol>
			<li>Luria-Bertani (LB) culture medium: Mix 10 g of tryptone, 5 g of yeast extract, and 10 g of sodium chloride in 1 L of ddH2O. Autoclave at 121&#176;C for 20 min.</li>
			<li>Lysis buffer: Mix 25 mM of HEPES (pH 7.4), 150 mM of KCl, 5 mM of MgCl2, 1x complete EDTA-free protease inhibitor cocktail, 10 mM of imidazole, and 0.5 mM of TCEP. Store at 4 &#176;C.</li>
			<li>Wash buffer: Mix 25 mM of HEPES (pH 7.4), 150 mM of KCl, 5 mM of MgCl2, 30 mM of imidazole, and 0.5mM of Tris(2-carboxyethyl)phosphine hydrochloride (TCEP). Store at 4 &#176;C.</li>
			<li>Elute buffer: Mix 25 mM of HEPES (pH 7.4), 150 mM of KCl, 5 mM of MgCl2, 150 mM of imidazole, and 0.5 mM of TCEP. Store at 4 &#176;C.</li>
			<li>Purification buffer: 25 mM of HEPES (pH 7.4), 150 mM of KCl, 5 mM of MgCl2, 0.5 mM of TCEP. Store at 4 &#176;C.</li>
		</ol>
		</li>
	</ol>
	</li>
	<li>Protein purification
	<ol>
		<li>Transfer the plasmids27 into Rosetta (DE3) (Biomed) E. coli cells using Bacterial transformation heat shock methods. Suspend the transferred cells in LB culture medium with a final concentration of 50 &#181;g/&#181;L kanamycin added.
		<ol>
			<li>Culture at 37 &#176;C until the absorbance of the culture medium at 600 nm is 0.6-0.8. Add isopropyl-&#946;-D-thiogalactoside (IPTG) at a final concentration of 50 &#181;M to induce the protein expression at 16 &#176;C for 20-24 h. 
			NOTE: The growth rate of bacterial or other microbial cell cultures can be monitored by measuring the optical density (absorbance) of the cell growth culture at 600 nm.</li>
		</ol>
		</li>
		<li>Harvest cells by centrifugation at 4000 &#215; g for 20 min at room temperature. Discard the supernatant.</li>
		<li>Resuspend the cell pellet with 30 mL of lysis buffer. Disrupt the cells three times using a high-pressure cell crusher.</li>
		<li>Clarify the lysates by centrifugation at 2,00,000 &#215; g for 1 h at 4 &#176;C using a ultracentrifuge rotor.</li>
		<li>Isolate the protein using a Ni-NTA column. Wash the Ni-NTA column with wash buffer for 5 column volumes. Elute protein with elution buffer for 1 column volume. Collect the eluate and concentrate using centrifugal filters to ~ 500 &#181;L.</li>
		<li>Further, purify the protein by size exclusion chromatography27. Collect the eluate (500 &#181;L per tube).</li>
		<li>Confirm the target protein by its molecular weight determined by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE)27. Collect the target protein for further modification.</li>
	</ol>
	</li>
</ol>
4. Protein biotinylation
<ol>
	<li>Preparation
	<ol>
		<li>Prepare the following stock solutions.
		<ol>
			<li>Dissolve Magnesium Acetate Tetrahydrate (MgOAc&#183;4H2O) in 25 mM of HEPES buffer (pH 7.7) at a concentration of 100 mM (10&#215;). Aliquot 100 &#181;L per tube and store at -20 &#176;C.</li>
			<li>Dissolve adenosine 5-triphosphate disodium salt (ATP) in ddH2O at a concentration of 100 mM (10&#215;). Adjust the pH to 7.4 using NaOH. Aliquot 100 &#181;L per tube and store at -80 &#176;C.</li>
			<li>Dissolve d-biotin in ddH2O at a concentration of 500 &#181;M. Aliquot 100 &#181;L per tube and store at -20 &#176;C.</li>
			<li>Dissolve streptavidin in ddH2O at a concentration of 1 mg/mL. Aliquot 20 &#181;L per tube and store at -20 &#176;C.</li>
		</ol>
		</li>
		<li>Protein biotinylation buffer: Add 25 mM of HEPES (pH 7.7), 200 mM of L-glutamic acid potassium, and 0.5 mM of TCEP. Store at 4 &#176;C.</li>
		<li>Purify BirA biotin ligase and store at -80 &#176;C.</li>
	</ol>
	</li>
	<li>Protein biotinylation
	<ol>
		<li>Change the protein buffer to the protein biotinylation buffer. For biotinylation, mix 100 &#181;L of 100 mM ATP, 100 &#181;L of 100 mM MgOAc, 100 &#181;L of 500 &#181;M d-biotin, 10 &#181;L of 100 &#181;M BirA biotin ligase, 50 &#181;M of protein to final volume of 1 mL and incubate overnight at 4 &#176;C.</li>
		<li>Remove free d-biotin using a 10 K MWCO centrifugal filter.</li>
		<li>Incubate 20 &#181;L biotinylated protein with 20 &#181;L 15 &#181;M streptavidin for 20 min on ice. Detect the efficiency of protein biotinylation by SDS-PAGE27.</li>
	</ol>
	</li>
</ol>
5. Protein fluorophore labeling
<ol>
	<li>Preparation
	<ol>
		<li>Dissolve fluorophores LD555 (donor) and LD655 (acceptor) in dimethyl sulfoxide (DMSO) to a final concentration of 5 mM. Store at -20 &#176;C.</li>
		<li>Prepare labeling buffer: Mix 25 mM of HEPES (pH 7.4), 150 mM of KCl, and 5 mM of MgCl2. Store at 4 &#176;C.</li>
		<li>Change the protein buffer to the labeling buffer.</li>
	</ol>
	</li>
	<li>Fluorophore labeling
	<ol>
		<li>For intramolecular smFRET assays, mix ATL1cyto-TK with LD555 and LD655 at a ratio of 1:1.2:1.2 to a final volume of 100 &#181;L (the total volume of fluorophores should not exceed 1 &#181;L), and incubate for 5 h at 4 &#176;C.</li>
		<li>For intermolecular smFRET experiments, incubate ATL1cyto-K or ATL1cyto-T with LD555 at a ratio of 1:1.2, and simultaneously incubate biotinylated ATL1cyto-K with LD655 at a ratio of 1:1.2, for 5 h at 4 &#176;C (same incubation volume as in 5.2.1). 
		NOTE: Co-incubation of protein and dyes allows flexibility in labeling positions for LD555 (donor) and LD655 (acceptor). The fact that LD555 is labeled to T51C or K400C does not affect the results of the FRET experiment. A schematic diagram of the donor being labeled at the T51C site is provided (Figure 1C,D).</li>
		<li>Remove excess free fluorophores using 7 K MWCO spin desalting columns27.</li>
		<li>Add TCEP to fluorophore-labeled proteins at a final concentration of 0.5 mM.</li>
		<li>Detect the labeling efficiency of proteins using a spectrophotometer. Measure protein absorbance at 280 nm, LD555 absorbance at 555 nm, and LD555 absorbance at 655 nm, and calculate the molar concentrations using their extinction coefficients, respectively. Protein labeling efficiency = (molar concentration of fluorescent dye/molar concentration of protein) x 100%.</li>
	</ol>
	</li>
</ol>
6. Single-molecule FRET imaging
<ol>
	<li>Preparation
	<ol>
		<li>Prepare the incubation buffer: Mix 25 mM of HEPES (pH 7.4), 150 mM of KCl, and 5 mM of MgCl2. Store at 4 &#176;C.</li>
		<li>Prepare 1 mg/mL streptavidin (dissolved in ddH2O) and 1 mg/mL biotinylated anti-His antibody for immobilization of proteins.</li>
		<li>Prepare 25 mM benzoic acid (pH 7.4; dissolved in the buffer as described in 7.1.1) and 100 mM of protocatechuate 3,4-dioxygenase (PCD) (dissolved in 40% glycerin solution) to minimize photobleaching during the smFRET experiments.</li>
		<li>Prepare 100 mM of Guanosine 5&#39;-diphosphate sodium salt (GDP), Guanosine 5&#39;-O-[gamma-thio]triphosphate (GTP&#947;S), and AlCl3 stock solutions dissolved in ddH2O. Prepare 1 M of NaF stock solution dissolved in ddH2O.</li>
		<li>Take out the vacuum-preserved coverslip and microscope slide, and carefully stick them together with customized double-sided tape on a clean bench. Install hoses and tips to form a microfluidic chamber with six channels27. 
		NOTE: Do not expose the modified coverslip to air for extended periods of time. Prepare the microfluidic chamber27 before starting the smFRET experiments.</li>
	</ol>
	</li>
	<li>Mapping correction 
	NOTE: During the experiment, fluorescence signals are to be collected from the donor and acceptor channels, respectively. Due to the use of microscopes, there may be slight deviations in the monitoring fields of the donor and acceptor channels. In order to reduce the deviation in the monitoring fields, mapping calibration must be performed before starting the experiment.
	<ol>
		<li>Mix 10% polystyrene particles (diameter 3 &#181;m) thoroughly, add 10 &#181;L of particles to a microscope slide (not modified), and cover with a coverslip (not modified).</li>
		<li>Select a field of view with as many polystyrene particles as possible. Capture a video under a bright field using total internal reflection fluorescence (TIRF) microscopy. 
		NOTE: A high numerical aperture oil immersion objective (100x) was used for high-resolution imaging. For smFRET experiments, signals were captured at 30 ms frame intervals using an EMCCD camera.</li>
		<li>Use a custom script to align the center location of the same polystyrene particle in the donor channel and acceptor channel and save the map file as a txt file. The custom MATLAB code used in this study is available at&#160;https://github.com/yangchenguang-1994/HMM-FRET).</li>
	</ol>
	</li>
	<li>Immobilizing the proteins in the chamber
	<ol>
		<li>For intermolecular smFRET experiments, mix LD555-labeled ATL1cyto-K or ATL1cyto-T and LD655-labeled ATL1cyto-K-biotin at a ratio of 1:1 with a final concentration of 1 mM of GTP&#947;S or GDP/AlF4-, and incubate for 1 h on ice to dimerize the protein.</li>
		<li>For ATL1cyto-T and ATL1cyto-K intermolecular smFRET experiments, mix LD555-labeled ATL1cyto-T and LD655-labeled ATL1cyto-K-biotin at a ratio of 1:1 with a final concentration of 1 mM of GTP&#947;S or GDP/AlF4-, and incubate for 1 h on ice to obtain dimerized proteins.</li>
		<li>For intramolecular smFRET assays, incubate LD555 and LD655-labeled ATL1cyto-TK with 1 mM of GDP for 1 h on ice before performing the smFRET experiments under GDP conditions.</li>
		<li>For intermolecular smFRET experiments, incubate 10 &#181;g/mL streptavidin (dilute with 200 &#181;L incubation buffer) for 10 min to immobilize proteins. For intramolecular smFRET assays, add 10 &#181;g/mL biotinylated anti-His antibody (diluted with 200 &#181;L incubation buffer) and incubate for 10 min to immobilize proteins. Flush the channel once with an incubation buffer before adding each solution.</li>
		<li>Dilute ATL1 dimer to ~0.3 nM or monomer to ~1 nM in 200 &#181;L incubation buffer, and incubate for 10 min to immobilize proteins in the channel. Flush the channel with incubation buffer twice. Add benzoic acid and PCD to the channel at a final concentration of 2.5 mM (diluted with 200 &#181;L buffer). 
		NOTE: To simulate the different stages of ATL1cyto molecules in the GTP hydrolysis process, nucleotide (GDP, GTP&#947;S, and GDP/AlF4-) was added at a final concentration of 1 mM to the buffers in each step after immobilizing the protein in the channel. GTP&#947;S addition mimics GTP binding, GDP/AlF4- addition mimics GTP hydrolysis, GDP addition mimics Pi release, and apo sate represents GDP release.</li>
	</ol>
	</li>
	<li>smFRET imaging
	<ol>
		<li>Adjust the focal plane. Use the coarse and fine focus knobs, adjust the focal plane to bring the sample into sharp focus. Here, an automated focus system was used to achieve the optimal focus.</li>
		<li>Set Up the Excitation Source. Use the 532 nm laser as the excitation light source. Adjust the laser power to a suitable level to excite the fluorophores. Here, a laser power of 20-40 mW was used.</li>
		<li>Connect the EMCCD camera to the microscope. Set the camera to record at a frame interval of 30 ms. Ensure the camera is set to capture images in 16-bit mode for high-resolution data.</li>
		<li>Record the movie, ensuring that the focus remains steady throughout the acquisition. Extend the 532nm laser irradiation time as long as possible until most of the fluorescent molecules are quenched when recording the fluorescence signals.</li>
		<li>Save the recorded movies in 16-bit TIFF format for further analysis.</li>
	</ol>
	</li>
</ol>
7. Data acquisition and analysis
<ol>
	<li>Load the map txt file for correction before extracting the FRET trajectories from the recorded movies. Select the FRET trajectories and save every FRET trajectory data in txt format for further analysis.</li>
	<li>Extract the data from the txt files and calculate the FRET values using homemade scripts in Matlab2022a. The custom MATLAB code is available at&#160;https://github.com/yangchenguang-1994/HMM-FRET. 
	NOTE: The FRET value is calculated using the equation ILD655/(&#947;ILD555+ILD655). ILD555 and ILD655 represent the LD555 and LD655 fluorescence intensities, respectively. &#947; is a parameter obtained by the formula &#947; = FA/FD. FA represents the decrement of acceptor intensity, and FD is the increment of donor intensity.</li>
	<li>Fit the smFRET data by GaussAmp in Origin 2022 to obtain the distribution histogram. 
	NOTE: For data with multiple FRET states (the data under GTP&#947;S condition in intermolecular smFRET or the data of intramolecular smFRET under GDP or Apo condition), multi-peak fitting can be used.</li>
</ol>

Monitoring GTPase Atlastin in Different Nucleotide Loading States

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<li>Byrnes, L. J., Sondermann, H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Structural+basis+for+the+nucleotide-dependent+dimerization+of+the+large+g+protein+atlastin-1%2Fspg3a%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">Structural basis for the nucleotide-dependent dimerization of the large g protein atlastin-1/spg3a.</a> Proc Natl Acad Sci U S A. 108 (6), 2216-2221 (2011).</li>
<li>Morin-Leisk, J., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((An+intramolecular+salt+bridge+drives+the+soluble+domain+of+gtp-bound+atlastin+into+the+postfusion+conformation%5BTitle%5D)+AND+%22J+Cell+Biol%22%5BJournal%5D)">An intramolecular salt bridge drives the soluble domain of gtp-bound atlastin into the postfusion conformation.</a> J Cell Biol. 195 (4), 605-615 (2011).</li>
<li>Pendin, D., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((GTP-dependent+packing+of+a+three-helix+bundle+is+required+for+atlastin-mediated+fusion%5BTitle%5D)+AND+%22Proc+Natl+Acad+Sci+U+S+A%22%5BJournal%5D)">GTP-dependent packing of a three-helix bundle is required for atlastin-mediated fusion.</a> Proc Natl Acad Sci U S A. 108 (39), 16283-16288 (2011).</li>
<li>Winsor, J., Hackney, D. D., Lee, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+crossover+conformational+shift+of+the+GTPase+atlastin+provides+the+energy+driving+ER+fusion%5BTitle%5D)+AND+%22J+Cell+Biol%22%5BJournal%5D)">The crossover conformational shift of the GTPase atlastin provides the energy driving ER fusion.</a> J Cell Biol. 216 (5), 1321-1335 (2017).</li>
<li>Yan, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Structures+of+the+yeast+dynamin-like+GTPase+sey1p+provide+insight+into+homotypic+er+fusion%5BTitle%5D)+AND+%22J+Cell+Biol%22%5BJournal%5D)">Structures of the yeast dynamin-like GTPase sey1p provide insight into homotypic er fusion.</a> J Cell Biol. 210 (6), 961-972 (2015).</li>
<li>Rizo, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Molecular+mechanisms+underlying+neurotransmitter+release%5BTitle%5D)+AND+%22Annu+Rev+Biophys%22%5BJournal%5D)">Molecular mechanisms underlying neurotransmitter release.</a> Annu Rev Biophys. 51, 377-408 (2022).</li>
<li>Gao, S., Hu, J. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mitochondrial+fusion%3A+The+machineries+in+and+out%5BTitle%5D)+AND+%22Trends+Cell+Biol%22%5BJournal%5D)">Mitochondrial fusion: The machineries in and out.</a> Trends Cell Biol. 31 (1), 62-74 (2021).</li>
<li>Kelly, C. M., Byrnes, L. J., Neela, N., Sondermann, H., O'donnell, J. P. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((The+hypervariable+region+of+atlastin-1+is+a+site+for+intrinsic+and+extrinsic+regulation%5BTitle%5D)+AND+%22J+Cell+Biol%22%5BJournal%5D)">The hypervariable region of atlastin-1 is a site for intrinsic and extrinsic regulation.</a> J Cell Biol. 220 (11), e202104128(2021).</li>
<li>Asher, W. B., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((GPCR-mediated+beta-arrestin+activation+deconvoluted+with+single-molecule+precision%5BTitle%5D)+AND+%22Cell%22%5BJournal%5D)">GPCR-mediated beta-arrestin activation deconvoluted with single-molecule precision.</a> Cell. 185 (10), 1661-1675.e16 (2022).</li>
<li>Li, M., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mechanism+of+DNA+translocation+underlying+chromatin+remodelling+by+snf2%5BTitle%5D)+AND+%22Nature%22%5BJournal%5D)">Mechanism of DNA translocation underlying chromatin remodelling by snf2.</a> Nature. 567 (7748), 409-413 (2019).</li>
<li>Chen, Y., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Conformational+dynamics+of+dynamin-like+MXA+revealed+by+single-molecule+fret%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">Conformational dynamics of dynamin-like MXA revealed by single-molecule fret.</a> Nat Commun. 8, 15744(2017).</li>
<li>Shi, L., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Dissecting+the+mechanism+of+atlastin-mediated+homotypic+membrane+fusion+at+the+single-molecule+level%5BTitle%5D)+AND+%22Nat+Commun%22%5BJournal%5D)">Dissecting the mechanism of atlastin-mediated homotypic membrane fusion at the single-molecule level.</a> Nat Commun. 15 (1), 2488(2024).</li>
<li>Guelly, C., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Targeted+high-throughput+sequencing+identifies+mutations+in+atlastin-1+as+a+cause+of+hereditary+sensory+neuropathy+type+i%5BTitle%5D)+AND+%22Am+J+Hum+Genet%22%5BJournal%5D)">Targeted high-throughput sequencing identifies mutations in atlastin-1 as a cause of hereditary sensory neuropathy type i.</a> Am J Hum Genet. 88 (1), 99-105 (2011).</li>
<li>Montagna, A., Vajente, N., Pendin, D., Daga, A. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((In+vivo+analysis+of+CRISPR%2FCas9+induced+atlastin+pathological+mutations+in+drosophila%5BTitle%5D)+AND+%22Front+Neurosci%22%5BJournal%5D)">In vivo analysis of CRISPR/Cas9 induced atlastin pathological mutations in drosophila.</a> Front Neurosci. 14, 547746(2020).</li>
<li>Ulengin, I., Park, J. J., Lee, T. H. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((ER+network+formation+and+membrane+fusion+by+atlastin1%2Fspg3a+disease+variants%5BTitle%5D)+AND+%22Mol+Biol+Cell%22%5BJournal%5D)">ER network formation and membrane fusion by atlastin1/spg3a disease variants.</a> Mol Biol Cell. 26 (9), 1616-1628 (2015).</li>
<li>Zhao, X., et al. <a target="_blank" href="http://www.ncbi.nlm.nih.gov/pubmed?term=((Mutations+in+a+newly+identified+GTPase+gene+cause+autosomal+dominant+hereditary+spastic+paraplegia%5BTitle%5D)+AND+%22Nat+Genet%22%5BJournal%5D)">Mutations in a newly identified GTPase gene cause autosomal dominant hereditary spastic paraplegia.</a> Nat Genet. 29 (3), 326-331 (2001).</li>
</ol>

The authors declare no conflicts of interest.

FRET technology is based on energy transfer between two fluorescent dyes (donor and acceptor). In the case of their proximity to each other (usually 1-10 nm), the excited donor transfers its energy to the acceptor, resulting in a decrease in the donor fluorescence intensity and an increase in the acceptor fluorescence intensity.
In smFRET experiments, the molecules are diluted to very low concentrations and immobilized on slides, and the FRET signals of individual molecules are detected by TIRF microscopy. Donor and acceptor fluorescent dyes are covalently attached to specific positions in molecules, and FRET efficiency is anti-correlated with the distance between fluorophores when the proteins undergo conformational changes or interact with other molecules. Therefore, the ratio of acceptor and donor fluorescence intensities can be measured to provide important information on the conformational dynamics, folding kinetics, and polymer formation and dissociation of individual biological macromolecules.
In the current assays, the cysteines in the protein was used to label the fluorophores with maleimide residues. However, using this method to label proteins requires mutation of the cysteine residues in the protein to other amino acids (e.g., alanine) and mutation of the amino acid that needs to be marked to cysteine. The introduction of multiple mutations may lead to changes in the protein structure and function. Therefore, before performing single-molecule experiments, it is necessary to analyze the impact of multiple mutations on the protein. In addition to the reaction between maleimide and cysteine, the same labeling can be achieved by incorporating unnatural amino acids into the target protein and using click chemical reactions to label fluorophores at specific sites on the protein. Moreover, the labeling efficiency of the protein can affect the data quality of smFRET.
During the smFRET imaging of ATL1cyto, it was found that the fluorophores labeling the protein had lower total fluorescence values, which may be caused by protein-induced fluorescence modulation (PIFQ). Based on this, parameter &#947; was introduced to correct the final FRET value. This phenomenon especially occurs in intermolecular smFRET, indicating that self-dimerizing protein molecules may be more prone to PIFQ. Therefore, when studying proteins that tend to form multimers, the PIFQ of fluorophore-labeled proteins should be considered.
The domain rearrangements of ATL1 coupled with GTP hydrolysis have been shown to be critical to its ability to mediate membrane fusion but, due to technical limitations, the conformational dynamics of ATL1 in different nucleotide-loading states are not available. The application of smFRET to studying the conformational changes in ATL1 provides a coherent picture of ATL function27. Furthermore, there has been increasing evidence that HSP, HSN, and other related neurodegenerative diseases are caused by defects in the ER homotypic fusion mediated by ATL6,10,28,29,30,31. Most disease mutations in ATL1 affect nucleotide binding, protein dimerization, or both. However, some mutations affect ATL function via an unclear mechanism. The smFRET assays contribute to comprehensively elucidating the pathological mechanism and proposing corresponding treatment strategies27. Further studies applying smFRET technology have the potential to understand the molecular mechanisms of other members of the dynamin superfamily.

In eukaryotic cells, the dynamin superfamily proteins mediate the remodeling of biological membranes, including membrane tubulation, fission, and fusion1,2,3. Mutations in these proteins lead to a variety of human diseases, such as neurodegenerative diseases. The members of the dynamin superfamily typically contain a GTPase domain, a middle domain consisting of helical bundles, a motif for membrane binding, and a GTPase effector domain (GED). One such dynamin-like GTPase is atlastin (ATL), which catalyzes homotypic membrane fusion of the endoplasmic reticulum (ER) to form a network4,5,6,7,8,9,10. There are three ATLs (ATL1-3) in mammals. The ATL molecule consists of an N-terminal cytosolic region (ATLcyto), including a GTPase domain and a three-helical middle domain (3HB), followed by two transmembrane regions, but lacks the typical GED. Alternatively, ATL contains a C-terminal tail that plays a crucial role in membrane fusion11,12,13.
Efficient membrane fusion has been reported to rely on the GTP hydrolysis-dependent domain rearrangement in ATLcyto14,15,16,17,18,19,20. However, compared to the SNARE complex, which uses folding energy from conformational changes to drive heterotypic membrane fusion21, the homotypic membrane fusion process mediated by ATL and other fusion-specific dynamin-like proteins22 remains elusive. Three crystal structures of human ATL1cyto have been determined in different nucleotide-loading conditions14,15,16,23. In the structures, the two 3HBs in the dimer point in different directions, but a comprehensive model explaining how the conformational dynamics of ATLcyto and its GTPase activity are coupled is still missing.
In the previous model, the monomeric ATL molecules form dimers across different membranes in a GTP-dependent manner to drive membrane fusion. Here, three strategies are described to apply single-molecule F&#246;rster resonance energy transfer (smFRET) to directly observe and accurately detect the behavior of individual ATL1cyto molecules. smFRET is a powerful technique extensively utilized to investigate conformational dynamics and interactions of individual biomolecules in real-time, such as GPCR-mediated &#946;-arrestin activation24, chromatin remodeling by Snf225, and domain rearrangements of dynamin-like protein MxA coupled with GTP hydrolysis cycles26. Since smFRET can only effectively detect changes in distance from ~3 nm to ~8 nm, both intermolecular and intramolecular smFRET experiments are required to comprehensively analyze the conformations of ATL1cyto for each nucleotide-loading state. We generated three constructs of ATL1cyto with all native cysteines substituted with alanines and K400 (ATL1cyto-K) or the T51/K400 pair (ATL1cyto-TK) mutated to cysteine for fluorophore labeling. The strategies for immobilizing the ATL1cyto dimer or monomer in a streptavidin-coated microfluidic chamber (Figure 1A) for intermolecular or intramolecular smFRET experiments are shown in Figure 1B-D. This method provides more details on the conformational dynamics of dynamin-like proteins in each step of GTP hydrolysis27.

<table><tbody><tr><td>10 K Centrifugal Filters</td><td>Amicon</td><td>UFC901096</td><td></td></tr><tr><td>3-Aminopropyltriethoxysilane (APTES)&amp;nbsp;</td><td>Sigma-Aldrich</td><td>440140</td><td></td></tr><tr><td>45 Ti rotor</td><td></td><td></td><td></td></tr><tr><td>532-nm laser</td><td>Olympus</td><td></td><td></td></tr><tr><td>640-nm laser</td><td>Olympus</td><td></td><td></td></tr><tr><td>7 K MWCO spin desalting columns</td><td>Thermo Scientific</td><td>89882</td><td></td></tr><tr><td>Adenosine 5-triphosphate disodium salt (ATP)</td><td>Roche</td><td>11140965001</td><td></td></tr><tr><td>Benzoic acid</td><td>Sigma-Aldrich</td><td>242381</td><td></td></tr><tr><td>Biotin-PEG-SVA-5000</td><td>Laysan Bio</td><td>170-124</td><td></td></tr><tr><td>Biotinylated anti His antibody</td><td>Bioss Antibodies</td><td>bs-0287R-bio</td><td></td></tr><tr><td>d-biotin</td><td>Sigma-Aldrich</td><td>B4501</td><td></td></tr><tr><td>EDTA-free protease inhibitor cocktail</td><td>Roche</td><td>11873580001</td><td></td></tr><tr><td>EMCCD camera</td><td>Andor</td><td>IX897</td><td></td></tr><tr><td>Guanosine 5&amp;prime;-diphosphate sodium salt (GDP)</td><td>Sigma-Aldrich</td><td>&amp;nbsp;G7127</td><td></td></tr><tr><td>Guanosine 5&#039;-O-[gamma-thio]triphosphate (GTP&amp;gamma;S)</td><td>Roche</td><td>10220647001</td><td></td></tr><tr><td>High numerical aperture oil immersion objective</td><td>Nikon</td><td></td><td>Nikon, 100x, N.A. 1.49, oil immersion</td></tr><tr><td>ImageJ</td><td>NIH</td><td></td><td>https://imagej.net/ij/</td></tr><tr><td>Isopropyl-&amp;Beta;-D-Thiogalactoside</td><td>Sigma-Aldrich</td><td>I5502</td><td></td></tr><tr><td>LD555-MAL</td><td>Lumidyne</td><td>4</td><td></td></tr><tr><td>LD655-MAL</td><td>Lumidyne</td><td>10</td><td></td></tr><tr><td>L-glutamic acid potassium</td><td>Sigma-Aldrich</td><td>G1501</td><td></td></tr><tr><td>Magnesium acetate tetrahydrate</td><td>Sigma-Aldrich</td><td>M0631</td><td></td></tr><tr><td>Matlab</td><td>The MathWorks, Natick, MA</td><td></td><td>https://www.mathworks.com/</td></tr><tr><td>Microscope cover glass</td><td>Fisherbrand</td><td>18834</td><td>&amp;nbsp;(24 mm &amp;times; 60 mm)</td></tr><tr><td>Microscope slides</td><td></td><td></td><td>Customized, (75 mm &amp;times; 26 mm &amp;times; 1 mm) with 6 pairs of 1.2 mm diameter through holes</td></tr><tr><td>mPEG-SVA-5000</td><td>Laysan Bio</td><td>170-106</td><td></td></tr><tr><td>Ni Sepharose 6 Fast Flow</td><td>Cytiva</td><td>17531802</td><td></td></tr><tr><td>Origin</td><td>Origin software</td><td></td><td>https://www.originlab.com/</td></tr><tr><td>Polystyrene particles</td><td>QDSphere</td><td>AG1265</td><td></td></tr><tr><td>Protocatechuate 3,4-dioxygenase</td><td>Sigma-Aldrich</td><td>P8279</td><td></td></tr><tr><td>Streptavidin</td><td>Sangon Biotech</td><td>A610492</td><td></td></tr><tr><td>Superdex 200 Increase (10/300 GL)</td><td>Cytiva</td><td>28990944</td><td></td></tr><tr><td>TIRF microscope</td><td>Nikon</td><td>Ti2</td><td></td></tr><tr><td>Tris(2-carboxyethyl)phosphine hydrochloride (TCEP)</td><td>Thermo</td><td>75259</td><td></td></tr></tbody></table>

single-molecule fret imaging for observing the conformational dynamics of dynamin-like gtpase atlastin

The rigid-body rotation of the three-helical middle domain (3HB) relative to the GTPase domain of dynamin-like protein atlastin (ATL) is a crucial driver of homotypic membrane fusion within the endoplasmic reticulum (ER). Disruptions in this process have been associated with hereditary spastic paraplegia (HSP), a neurodegenerative disorder. Structural and biochemical studies suggest that the conformational changes in ATL are linked to GTP hydrolysis, but real-time visualization of these conformational dynamics during the GTP hydrolysis cycle remains challenging. To better understand the mechanical mechanisms behind ATL function, single-molecule F&#246;rster resonance energy transfer (smFRET) was utilized. Three specific strategies were employed to immobilize the N-terminal cytosolic region of human ATL1 (ATL1cyto) in a streptavidin-coated microfluidic chamber, facilitating the application of intramolecular and intermolecular smFRET imaging. This allowed precise monitoring of protein conformations in various nucleotide-loading states, providing direct insights into individual molecular behaviors. This method can be applied to study other mechanochemical proteins as well.

The smFRET experiments were performed on a TIRF microscope by capturing the fluorescence intensity of fluorophores every 30 ms. Representative images of LD555 (donor) and/or LD655 (acceptor)-labeled ATL1cyto molecules in the absence or presence of different nucleotides are shown in Figure 2A-C. The fluorescence intensities of the two fluorophores on individual particles were recorded, and hundreds of FRET trajectories exhibiting typical single-molecule features were extracted. The representative traces are shown in Figure 2D-F. In the absence or presence of GDP, ATL1cyto monomers displayed dynamic FRET trajectories with four FRET states (Figure 2D), indicating that the GTPase domain-3HB region of ATL1 is flexible in these states, but the GDP-bound ATL1cyto prefers the close conformation and the GDP release alters the conformational preference of ATL1cyto towards an open conformation (Figure 1B and Figure 2D)27. Upon GTP binding and in the transition state of GTP hydrolysis, ATL1cyto dimers exhibited stable FRET trajectories with one low-FRET state (Figure 2E), demonstrating that only the crossover dimer of ATLcyto forms (Figure 1C and Figure 2E)27. In the intermolecular smFRET experiments, only a high-FRET state was observed with GDP/AlF4-, whereas GTP&#947;S-bound ATL1cyto exhibited an additional low-FRET peak (Figure 2F)27. These results suggest that the crossover dimer of ATL1cyto is tightened by GTP hydrolysis (Figure 1D and Figure 2F)27.
<img alt="Figure 1" class="xfigimg" src="/files/ftp_upload/67263/67263fig01.jpg" /> 
Figure 1: Schematic of the smFRET experiments. (A) Design of a microfluidic chamber. (B-D) Strategies for smFRET assays. (B) LD555/LD655-labeled ATL1cyto-TK monomers (intramolecular smFRET assays for ATL1cyto monomer). (C) LD555/LD655-labeled ATL1cyto-TK and biotinylated ATL1cyto-K dimers (intramolecular smFRET assays for ATL1cyto dimer). (D) LD555-labeled ATL1cyto-K and biotinylated LD655-labeled ATL1cyto-K dimers (intermolecular smFRET assays for ATL1cyto dimer). The green spheres represent the LD555 (donor) labeling, and the red spheres represent the LD655 (acceptor) labeling. Nucleotides are shown as orange sticks, and magnesium ions are shown as yellow spheres. Ribbon representations of the crystal structures of ATL1cyto dimers or monomers in Form 1 (left, PDB code 3QOF), Form 2 (middle, PDB code 3QNU), and Form 3 (right, PDB code 4IDN) conformations rendered in PyMOL. The figure is adapted from Shi et al.27. <a href="https://www.jove.com/files/ftp_upload/67263/67263fig01large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>
<img alt="Figure 2" class="xfigimg" src="/files/ftp_upload/67263/67263fig02.jpg" /> 
Figure 2: Representative smFRET results. (A-C) Representative TIRF images of immobilized particles for the three strategies. Scale bar: 10 &#181;m. (D-F) Representative fluorescence and smFRET trajectories of the samples and histograms of smFRET data. LD555 (the donor) is shown in green, LD655 (the acceptor) is shown in red, and FRET is displayed in blue. The smFRET values with the number of molecules (Nm) displayed were compiled into a conformation-population histogram and fitted into GaussAmp distributions. Each bar height represents the normalized count (%). The length of the error bar represents the normalized SD of a Poisson distribution from the count. (A,D) Intramolecular smFRET results for ATL1cyto monomer. (B,E) Intramolecular smFRET results for ATL1cyto dimer. (C,F) Intermolecular smFRET results for ATL1cyto dimer. The figure is adapted from Shi et al.27. <a href="https://www.jove.com/files/ftp_upload/67263/67263fig02large.jpg" target="_blank">Please click here to view a larger version of this figure.</a>

Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

The functions of dynamin superfamily proteins depend on conformational changes coupled with GTP hydrolysis. A system is described using the single-molecule FRET (smFRET) technique to monitor the conformational dynamics of dynamin-like GTPase atlastin in different nucleotide-loading states.

The rigid-body rotation of the three-helical middle domain (3HB) relative to the GTPase domain of dynamin-like protein atlastin (ATL) is a crucial driver ...

single-molecule-fret-imaging-for-observing-conformational-dynamics

Universidad Científica del Sur

Research

JoVE Journal

Biochemistry

388 Views.  Nankai University. The functions of dynamin superfamily proteins depend on conformational changes coupled with GTP hydrolysis. A system is described using the single-molecule FRET (smFRET) technique to monitor the conformational dynamics of dynamin-like GTPase atlastin in different nucleotide-loading states.

Video: Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

Magnetic Tweezers in a Microplate Format

Mechanobiology describes how the physical forces and mechanical properties of biological material contribute to physiology and disease. Typically, these approaches are limited single-molecule methods, which restricts their availability. To address this need, a microplate assay was developed that enables mechanical manipulation while performing standard biochemical assays. This is achieved using magnets incorporated into a microplate lid to create multiple magnetic tweezers. In this format, force is exerted across biomolecules connected to paramagnetic beads, equivalent to a typical magnetic tweezer. The study demonstrates the application of this tool with FRET-based assays to monitor protein conformations. However, this approach is widely applicable to different biological systems ranging from measuring enzymatic activity through to the activation of signaling pathways in live cells.

Here, we describe the use of a novel microplate assay to enable mechanical manipulation of biomolecules while performing ensemble biochemical assays. This is achieved using a microplate lid modified with magnets to create multiple static magnetic tweezers across the microplate.

Mechanobiology describes how the physical forces and mechanical properties of biological material contribute to physiology and disease. Typically, these ...

Using Microfluidics and Fluorescence Microscopy to Study the Assembly Dynamics of Single Actin Filaments and Bundles

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor individual reactions live in well-controlled conditions. To do so, live single-filament experiments have emerged over the past 20 years, mostly using total internal reflection fluorescence (TIRF) microscopy, and have provided a trove of key results. In 2011, in order to further expand the possibilities of these experiments and to avoid recurring problematic artifacts, we introduced simple microfluidics in these assays. This study details our basic protocol, where individual actin filaments are anchored by one end to the passivated coverslip surface, align with the flow, and can be successively exposed to different protein solutions. We also present the protocols for specific applications and explain how controlled mechanical forces can be applied, thanks to the viscous drag of the flowing solution. We highlight the technical caveats of these experiments and briefly present possible developments based on this technique. These protocols and explanations, along with today&#39;s availability of easy-to-use microfluidics equipment, should allow non-specialists to implement this assay in their labs.

We present protocols for simple actin filament microfluidic assays, in combination with fluorescence microscopy, that allow one to accurately monitor individual actin filaments in real-time while sequentially exposing them to different protein solutions.

In order to decipher the complex molecular mechanisms that regulate the assembly and disassembly of actin filaments, it is a great asset to monitor ...

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence (TIRF) Microscopy

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and regulated by different suites of binding proteins unique for each polymer. Many studies now demonstrate that the dynamics of both cytoskeletal polymers are intertwined and that this crosstalk is required for most cellular behaviors. A number of proteins involved in actin-microtubule interactions have already been identified (i.e., Tau, MACF, GAS, formins, and more) and are well characterized with regard to either actin or microtubules alone. However, relatively few studies showed assays of actin-microtubule coordination with dynamic versions of both polymers. This may occlude emergent linking mechanisms between actin and microtubules. Here, a total internal reflection fluorescence (TIRF) microscopy-based in vitro reconstitution technique permits the visualization of both actin and microtubule dynamics from the one biochemical reaction. This technique preserves the polymerization dynamics of either actin filament or microtubules individually or in the presence of the other polymer. Commercially available Tau protein is used to demonstrate how actin-microtubule behaviors change in the presence of a classic cytoskeletal crosslinking protein. This method can provide reliable functional and mechanistic insights into how individual regulatory proteins coordinate actin-microtubule dynamics at a resolution of single filaments or higher-order complexes.

Visualizing Actin and Microtubule Coupling Dynamics In Vitro by Total Internal Reflection Fluorescence TIRF Microscopy

This protocol is a guide for visualizing dynamic actin and microtubules using an in vitro total internal fluorescence (TIRF) microscopy assay.

Traditionally, the actin and microtubule cytoskeletons have been studied as separate entities, restricted to specific cellular regions or processes, and ...

Imaging the Motion of Fully Reconstituted Cdc45/Mcm2-7/GINS (CMG)

Hybrid Ensemble and Single-molecule Assay to Image the Motion of Fully Reconstituted CMG

Eukaryotes have one replicative helicase known as CMG, which centrally organizes and drives the replisome, and leads the way at the front of replication forks. Obtaining a deep mechanistic understanding of the dynamics of CMG is critical to elucidating how cells achieve the enormous task of efficiently and accurately replicating their entire genome once per cell cycle. Single-molecule techniques are uniquely suited to quantify the dynamics of CMG due to their unparalleled temporal and spatial resolution. Nevertheless, single-molecule studies of CMG motion have thus far relied on pre-formed CMG purified from cells as a complex, which precludes the study of the steps leading up to its activation. Here, we describe a hybrid ensemble and single-molecule assay that allowed imaging at the single-molecule level of the motion of fluorescently labeled CMG after fully reconstituting its assembly and activation from 36 different purified S. cerevisiae polypeptides. This assay relies on the double functionalization of the ends of a linear DNA substrate with two orthogonal attachment moieties, and can be adapted to study similarly complex DNA-processing mechanisms at the single-molecule level.

Author Spotlight: Investigating the Motion Dynamics of the Eukaryotic Replisome Components at the Single-Molecule Level

We report a hybrid ensemble and single-molecule assay to directly image and quantify the motion of fluorescently labeled, fully reconstituted Cdc45/Mcm2-7/GINS (CMG) helicase on linear DNA molecules held in place in an optical trap.

Eukaryotes have one replicative helicase known as CMG, which centrally organizes and drives the replisome, and leads the way at the front of replication ...

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. Single-molecule imaging is gaining momentum as it provides exceptional access to the visualization of individual molecules in living cells. Here, we describe a technique that we developed to perform single-particle tracking photo-activated localization microscopy (sptPALM) in Drosophila larvae. Synaptic communication relies on key presynaptic proteins that act by docking, priming, and promoting the fusion of neurotransmitter-containing vesicles with the plasma membrane. A range of protein-protein and protein-lipid interactions tightly regulates these processes and the presynaptic proteins therefore exhibit changes in mobility associated with each of these key events. Investigating how mobility of these proteins correlates with their physiological function in an intact live animal is essential to understanding their precise mechanism of action. Extracting protein mobility with high resolution in vivo requires overcoming limitations such as optical transparency, accessibility, and penetration depth. We describe how photoconvertible fluorescent proteins tagged to the presynaptic protein Syntaxin-1A can be visualized via slight oblique illumination and tracked at the motor nerve terminal or along the motor neuron axon of the third instar Drosophila larva.

In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

Here we illustrate how single molecule photo-activated localization microscopy can be carried out on the motor nerve terminal of a live Drosophila melanogaster third instar larva.

An increasing number of super-resolution microscopy techniques are helping to uncover the mechanisms that govern the nanoscale cellular world. ...

Neuroscience

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This feature is attributed to the mechanical and dynamic properties of the cell cytoskeleton, notably, the actomyosin cytoskeleton, which is an active gel of polar actin filaments, myosin motors, and accessory proteins that exhibit intrinsic contraction properties. The usually accepted view is that the cytoskeleton behaves as a viscoelastic material. However, this model cannot always explain the experimental results, which are more consistent with a picture describing the cytoskeleton as a poroelastic active material-an elastic network embedded with cytosol. Contractility gradients generated by the myosin motors drive the flow of the cytosol across the gel pores, which infers that the mechanics of the cytoskeleton and the cytosol are tightly coupled. One main feature of poroelasticity is the diffusive relaxation of stresses in the network, characterized by an effective diffusion constant that depends on the gel elastic modulus, porosity, and cytosol (solvent) viscosity. As cells have many ways to regulate their structure and material properties, our current understanding of how cytoskeleton mechanics and cytosol flow dynamics are coupled remains poorly understood. Here, an in vitro reconstitution approach is employed to characterize the material properties of poroelastic actomyosin gels as a model system for the cell cytoskeleton. Gel contraction is driven by myosin motor contractility, which leads to the emergence of a flow of the penetrating solvent. The paper describes how to prepare these gels and run experiments. We also discuss how to measure and analyze the solvent flow and gel contraction both at the local and global scales. The various scaling relations used for data quantification are given. Finally, the experimental challenges and common pitfalls are discussed, including their relevance to cell cytoskeleton mechanics.

The Mechanics of Poro-Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

In this work, an in vitro reconstitution approach is employed to study the poroelasticity of actomyosin gels under controlled conditions. The dynamics of the actomyosin gel and the embedded solvent are quantified, through which the network poroelasticity is demonstrated. We also discuss the experimental challenges, common pitfalls, and relevance to cell cytoskeleton mechanics.

Cells can actively change their shapes and become motile, a property that depends on their ability to actively reorganize their internal structure. This ...

Bioengineering

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in the x-y plane. Here a summary of the FIONA technique is reported and examples of research that have been performed using FIONA are briefly described. First, how to set up the required equipment for FIONA experiments, i.e., a total internal reflection fluorescence microscopy (TIRFM), with details on aligning the optics, is described. Then how to carry out a simple FIONA experiment on localizing immobilized Cy3-DNA single molecules using appropriate protocols, followed by the use of FIONA to measure the 36 nm step size of a single truncated myosin Va motor labeled with a quantum dot, is illustrated. Lastly, recent effort to extend the application of FIONA to thick samples is reported. It is shown that, using a water immersion objective and quantum dots soaked deep in sol-gels and rabbit eye corneas (&#62;200 &#181;m), localization precision of 2-3 nm can be achieved.

Fluorescence Imaging with One-nanometer Accuracy FIONA

Single fluorophores can be localized with nanometer precision using FIONA. Here a summary of the FIONA technique is reported, and how to carry out FIONA experiments is described.

Fluorescence imaging with one-nanometer accuracy (FIONA) is a simple but useful technique for localizing single fluorophores with nanometer precision in ...

Biology

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how fluorescent recovery after photobleaching (FRAP) and photoactivation techniques can be used to study the spatiotemporal dynamics of proteins in subcellular locations. We also show how these techniques enable straightforward determination of various parameters linked to actin cytoskeletal regulation and cell motility. Moreover, the microinjection of cells is additionally described as an alternative treatment (potentially preceding or complementing the aforementioned photomanipulation techniques) to trigger instantaneous effects of translocated proteins on cell morphology and function. Micromanipulation such as protein injection or local application of plasma membrane-permeable drugs or cytoskeletal inhibitors can serve as powerful tool to record immediate consequences of a given treatment on cell behavior at the single cell and subcellular level. This is exemplified here by immediate induction of lamellipodial cell edge protrusion by the injection of recombinant Rac1 protein, as established a quarter-century ago. In addition, we provide a protocol for determining the turnover of enhanced green fluorescent protein (EGFP)-VASP, an actin filament polymerase prominently accumulating at lamellipodial tips of B16-F1 cells, employing FRAP and including associated data analysis and curve fitting. We also present guidelines for estimating the rates of lamellipodial actin network polymerization, as exemplified by cells expressing EGFP-tagged &#946;-actin. Finally, instructions are given for how to investigate the rates of actin monomer mobility within the cell cytoplasm, followed by actin incorporation at sites of rapid filament assembly, such as the tips of protruding lamellipodia, using photoactivation approaches. None of these protocols is&#160;restricted to components or regulators of the actin cytoskeleton, but can easily be extended to explore in analogous fashion the spatiotemporal dynamics and function of proteins in various different subcellular structures or functional contexts.

We describe how micro- and photomanipulation techniques such as FRAP and photoactivation enable the determination of motility parameters and the spatiotemporal dynamics of proteins within migrating cells. Experimental readouts include subcellular dynamics and turnover of motility regulators or of the underlying actin cytoskeleton.

Examining the spatiotemporal dynamics of proteins can reveal their functional importance in various contexts. In this article, it is discussed how ...

Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy

Ultrafast force-clamp spectroscopy (UFFCS) is a single molecule technique based on laser tweezers that allows the investigation of the chemomechanics of both conventional and unconventional myosins under load with unprecedented time resolution. In particular, the possibility to probe myosin motors under constant force right after the actin-myosin bond formation, together with the high rate of the force feedback (200 kHz), has shown UFFCS to be a valuable tool to study the load dependence of fast dynamics such as the myosin working stroke. Moreover, UFFCS enables the study of how processive and non-processive myosin-actin interactions are influenced by the intensity and direction of the applied force.
By following this protocol, it will be possible to perform ultrafast force-clamp experiments on processive myosin-5 motors and on a variety of unconventional myosins. By some adjustments, the protocol could also be easily extended to the study of other classes of processive motors such as kinesins and dyneins. The protocol includes all the necessary steps, from the setup of the experimental apparatus to sample preparation, calibration procedures, data acquisition and analysis.

Presented here is a comprehensive protocol to perform ultrafast force-clamp experiments on processive myosin-5 motors, which could be easily extended to the study of other classes of processive motors. The protocol details all the necessary steps, from the setup of the experimental apparatus to sample preparation, data acquisition and analysis.

Ultrafast force-clamp spectroscopy (UFFCS) is a single molecule technique based on laser tweezers that allows the investigation of the chemomechanics of ...

Single-Molecule Imaging to Visualize Fusion Protein Condensates Assembly on DNA Curtains

To image the EWS-FLI1 condensation formation on DNA curtains, open the imaging software and find and mark the positions of the nine zig-zag patterns under bright-field. Then, turn on the flow at 0.2 milliliters per minute to stain the DNA with double-stranded DNA dye for 10 minutes.
Next, dilute the mCherry-EWS-FLI1 protein with the imaging buffer at a concentration of 100 nanomoles and 100 microliters. Then, load the protein sample through the valve with a 100-microliter glass syringe, and change the flow rate to 0.4 milliliters per minute.
After turning on the 488-nanometer laser, pre-scan each region to check the DNA distribution state, and select the region in which the DNA molecules distribute evenly. Then, set the laser power to 10% for the 488-nanometer laser and 20% for the 561-nanometer laser, and using the power meter, measure the real laser power near the prism.
Start acquiring images at two-second intervals with both 488 and 561-nanometer lasers simultaneously. Then, change the valve from the manual mode to injection mode to let the imaging buffer flush the protein sample into the flow cell after 60 seconds.
To remove the free EWS-FLI1, keep washing the flow cell with the imaging buffer for five minutes with only the 561-nanometer laser switched on. Then, stop the flow and incubate at 37 degrees Celsius for 10 minutes.
After 10 minutes, turn on the flow at 0.4 milliliters per minute to let the DNA extend, and acquire images at two-second intervals between different frames to obtain high-throughput data of EWS-FLI1 condensate formation.

Single-Molecule FRET Imaging for Observing the Conformational Dynamics of Dynamin-Like GTPase Atlastin

Summary

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In Vivo Single-Molecule Tracking at the Drosophila Presynaptic Motor Nerve Terminal

The Mechanics of (Poro-)Elastic Contractile Actomyosin Networks As a Model System of the Cell Cytoskeleton

Fluorescence Imaging with One-nanometer Accuracy (FIONA)

Micromanipulation Techniques Allowing Analysis of Morphogenetic Dynamics and Turnover of Cytoskeletal Regulators

Dissecting Mechanoenzymatic Properties of Processive Myosins with Ultrafast Force-Clamp Spectroscopy

Single-Molecule Imaging to Visualize Fusion Protein Condensates Assembly on DNA Curtains